Introduction
The term “plastics” denotes polymers, compounds characterized by their high molecular weight, derived from either natural or synthetic sources. These materials, categorized as high polymer substances, have been extensively utilized since the early 20th century [1]. Among them, there are various polymers such as polyethylene, polypropylene, and polystyrene, often enhanced with additives to augment their properties [2]. Plastic particles fall into two main groups: primary particles found in manufactured items, and secondary particles formed during the degradation of packaging and textiles. Through processes like photo-degradation and bio-degradation, plastics can fragment into different sizes, such as nanoplastics (≤0.1 μm), microplastics (MPs) (<5 mm), mesoplastics (0.5–5 cm), macroplastics (5–50 cm), and megaplastics (>50 cm) [3]. Notably, MPs, under 5 mm in diameter, come in various forms such as beads, fibers, and fragments, distinguished by their size, shape, and composition. Nanoplastics, however, encompass materials of natural, incidental, or synthetic origin, existing either as discrete particles or agglomerates held together by weak forces. The European Food Safety Authority defines them as particles ranging from 1 to 100 nm in size. Over time, plastics can degrade, breaking down into smaller particles, including MPs and nanoplastics [4]. Nanoplastics, potentially harmful to human health, can enter the food chain through contaminated food and water sources. The widespread use of plastics across numerous industries has led to their ubiquitous presence in modern goods, contributing to pollution in aquatic, terrestrial, and atmospheric environments globally [5]. Inadequate recycling efforts and a lack of regulations on plastic waste exacerbate this pollution. Plastic, including both nano and MPs, has infiltrated various ecosystems, from oceans and seas to rivers, lakes, and even Arctic sea ice [5]. Over the past six decades, global plastic production has exceeded 6.3 billion tons. Only about 9% of this has been recycled into secondary materials, while 12% has been incinerated for energy production [6]. The rate of decomposition of a 1-mm-thick plastic plate varies depending on its chemical composition and environmental conditions, leading to a prolonged disintegration process [7].
In recent years, mounting evidence has emerged linking MPs to immune system damage. These minuscule plastic particles have been shown to potentially harm immune cells and disrupt their normal functions. They can induce inflammation, impede communication between immune cells, and weaken the body’s defenses against infections and other health issues. Paul-Pont et al. [8] observed increased production of reactive oxygen species (ROS) and mortality among hemocytes in mussels exposed to polystyrene MPs (2 and 6 μm) at 32 μg/l for 7 days. Green et al. [9] found that blue mussels exposed to polyethylene MPs (0.48 to 316 μm) in drinking water for 52 days elevated levels of immune-responsive proteins. Alternatively, Revel et al. [10] explored the impacts of polyethylene and polypropylene MPs on ragworms over a 10-day ingestion period at concentrations ranging from 10 to 100 μg/l. They observed no significant alterations in phenoloxidase, acid phosphatase, or phagocytosis. Similarly, Revel et al. [11] found no discernible effects on DNA damage, acid phosphatase activity, or ROS production in Pacific oysters exposed to polyethylene and polypropylene fragments (<400 μm) in drinking water for 10 days, across doses of 0.008, 10, and 100 μg/l. In a recent study by Murano et al. [12], the effects of MP exposure on sea urchins were examined. The study focused on the influence of drinking water contaminated with various concentrations of MPs on sea urchins immune cell coelomocyte protein profiles. Abnormalities were detected in metabolite interconversion enzymes in cells exposed to MPs, indicating significant alterations in metabolic processes.
The research endeavors to enhance our comprehension by delving into the impact of MPs on the embryonic microenvironment. Specifically, it examines biochemical alterations in amniotic fluid and the modulation of immune genes. These findings added valuable insights to our existing knowledge. Our study focuses on the nuanced effects of MPs using the chick embryo model, chosen for its physiological resemblance to humans and its effectiveness in assessing developmental toxicity. We aim to deepen our understanding of the risks associated with MPs exposure during crucial embryonic stages. By exploring effects on the immune system and biochemical processes, we aim to grasp the intricate ways in which MPs influence embryonic development. Analyzing changes in amniotic fluid is crucial as it offers insights into how MPs impact the embryo’s microenvironment and immediate physiological responses [13]. This understanding is vital for evaluating the potential hazards and developmental consequences of MPs exposure. Our findings shed light on the fundamental mechanisms and risks associated with MPs, unveiling their effects on embryonic development and broader implications for avian health. By highlighting these crucial findings, our study aids informed decision-making and promotes responsible usage of MPs in various industrial and consumer applications.
Materials and Methods
MPs
Polystyrene MPs (PS-MPs) with a diameter of 4.8–5 μm were sourced from Cospheric LLC, USA. Their size renders them highly resistant to organic solvents when in dry powder form. The choice of a 5 μm diameter was intentional, aimed at facilitating laboratory research without interference from nanoparticles. Before commencing testing, the MP particles underwent a 30-minutes probe sonication process utilizing an ultrasonic cleaner (Model: TUC-32, China). Subsequently, the PS-MPs were meticulously prepared to achieve the required concentration by mixing them with distilled water (pH value of 6.9 ± 0.1).
Egg collection and dosing
Fertile chicken eggs sourced from the Sonali crossbreed (Gallus gallus domesticus), with an average weight of 50 ± 1.5 g, were acquired from the Government Poultry Farm situated in Rangpur-5400. These eggs were stored at 12°C for 48 hours before incubation. All in-ovo experimental methods strictly followed established protocols in our laboratory [14]. Ninety eggs were utilized, evenly distributed into three groups of 30 eggs each. The groups were labeled as follows: Group C (Control Group): 30 eggs injected with 0.3 ml saline solution, serving as a control group. Group A: 30 eggs treated with MPs at a dose of 150 µg/ml. Group B: 30 eggs receiving MPs at a dose of 300 µg/ml. After 48 hours of incubation, test samples (0.3 ml/egg) were administered into the air sac using sterile 1 ml tuberculin syringes. Subsequently, the eggs underwent an additional 18-days incubation period following procedures outlined by Patel et al. [15]. A mechanical system was employed to regulate temperature, humidity, and forced air ventilation throughout the incubation process. Specialized holders were used to rotate the eggs once every two hours, totaling twelve rotations daily, to facilitate optimal development. Embryo viability was monitored daily through candling across the three treatment groups and the control group, with incubation conditions maintained at 37.5°C and 60% humidity. Temperature fluctuations were closely monitored in all groups, ensuring stability with the incubator’s automatic temperature management system maintaining a consistent temperature of 37.5°C throughout the trial.
Ethical statement
The experimental protocol for overseeing chick embryo experiments was officially approved by the Ethical Committee of the Institute of Research and Training, under reference number (HSTU/IRT/2023/4357).
Specimen collection
The research was conducted over a period from June to December 2023. On the 18th day following incubation, five embryos were sampled at each designated time point for the purpose of evaluating biochemical parameters, as well as conducting histopathological and gene expression analyses. Concurrently, amniotic fluid was extracted, centrifuged, and analyzed for biochemical markers, specifically the activity levels of enzymes such as alkaline phosphatase (ALP), aspartate aminotransferase (AST), and alanine aminotransferase (ALT), in addition to urea and creatinine concentrations. On each collection day, the embryos were washed with saline and humanely euthanized by cooling. An incision was then made at the air sac end of each egg for further examination. Tissues from the caecal tonsil and bursa of Fabricius were carefully extracted on the 18th day of incubation. These samples were subsequently fixed in 10% neutral buffered formalin for histological assessments, and the caecal tonsil tissues were also preserved in Ribonucleic Acid (RNA) later for further gene expression analysis.
Histopathological studies
Formalin was removed from the tissues, which were then subjected to a series of alcohol dehydrations at gradual levels of 70, 80, 90, 95, and ultimately 100% for one hour at least in each level according to the protocol described by Islam et al. [14]. Following this procedure, the tissues were first soaked in xylene-1 for 90 minutes, then in xylene-2 for another 90 minutes. After xylene treatment, tissues were immersed in liquid paraffin at 60 °C for 90 minutes, then cooled and set into paraffin blocks. These blocks were sliced into 6 µm sections using a LEICA RM2125 RTS microtome. The sections were stretched on water at 45°C, mounted on clean, oil-free glass slides, and dried in an oven at 62°C for 20 minutes. Staining was performed with hematoxylin and eosin (H&E), followed by sealing the slides with Canada balsam, adhering to methods recommended by Drury et al. [16]. High-resolution images of the tissues were captured using an Amscope (MA500) camera attached to a Richter Ptica U-2T microscope at magnifications of 10×, 40×, and 100×. Image J software was utilized for the analysis of lymphocytes in the caecal tonsils, following the cell counting approach outlined by Curvo et al. [17] and employing Islam et al. [14] specified field length for consistent quantification. For each experimental condition, five slides were examined to calculate the average of cell counts.
Analysis of amniotic fluid biochemical parameters
On the 18th day of incubation, we analyzed the biochemical properties of amniotic fluid utilizing an 18-gauge needle to extract samples. Amniotic fluid was obtained from five eggs per experimental group. Following the careful removal of the air sac and membranes of the eggshell, each sample of amniotic fluid was subjected to centrifugation at 3,000 g for 15 minutes, isolating the supernatant for further testing. Biochemical analysis was conducted using a photometer (Siemens Advia 1800, Germany), targeting the detection of biochemical markers including the concentration of urea and creatinine as well as the activity of ALP, AST, and ALT.
Assessment of immune gene expression
Expression levels of AvBD9, AvBD10, and IL6 were evaluated using quantitative polymerase chain reaction (qPCR), with ACTB (β-actin) serving as the control gene for normalization purposes, as reported by Laptev et al. [18]. β-actin, chosen for its consistent expression among the samples, provided the cycle threshold (CT) values essential for standardizing gene expression data. To prepare for RNA extraction, the caecal tonsils from 18-day-old chick embryos were pulverized and mixed under liquid nitrogen. RNA was then isolated using the Monarch® Total RNA Miniprep Kit (Cat No. T2010S, New England Biolsabs Inc.), adapting the standard protocol slightly by incorporating liquid nitrogen to ensure thorough homogenization of the samples. The extraction was performed in triplicate for each group. Following RNA extraction, the ProtoScript® II First Strand cDNA Synthesis Kit (Cat No: E6560S, New England Biolabs Inc.) was employed to synthesize cDNA. The NANODROP ONE (Thermo Scientific, USA) was utilized to assess the purity and concentration of the RNA. For the qPCR, Luna Universal qPCR Master Mix (Cat No: M3003S, New England Biolabs Inc.) and the qTOWER3G real-time polymerase chain reaction (PCR) system (Analytik Jena) were used. The thermal cycling conditions were set to initial denaturation at 95°C for 2 minutes, followed by cycles of denaturation at 95°C for 5 seconds, annealing at 62°C for 30 seconds, and extension at 72°C for 30 seconds. The specific primers and expected product sizes for each gene were as follows: AvBD9 had a product size of 131 bp with forward primer AACACCGTCAGGCATCTTCACA and reverse primer CGTCTTCTTGGCTGTAAGCTGGA; AvBD10 had a product size of 102 bp with forward primer GCTCTTCGCTGTTCTCCTCT and reverse primer CCCAGAGATGGTGAAGGTG; IL6 had a product size of 78 bp with forward primer AGGACGAGATGTGCAAGAAGTTC and reverse primer TTGGGCAGGTTGAGGTTGTT; and the housekeeping gene ACTB (β-actin) had a product size of 86 bp with forward primer ATTGTCCACCGCAAATGCTTC and reverse primer AAATAAAGCCATGCCAATCTCGTC [18].
Statistical analysis
This research utilized SPSS version 16, a statistical software developed by SPSS, Inc., to perform data analysis. Initially, the data underwent a normality test to examine the distribution patterns of the variables. Subsequently, the study employed a one way analysis of variance (ANOVA) to analyze the significance levels of the data. The purpose of the ANOVA was to identify any statistically significant effects of treatment. A result was deemed significant if it met the 95% confidence threshold (p < 0.05), and highly significant if it met the 99% confidence threshold (p ≤ 0.01). In cases where the ANOVA revealed a significant effect of treatment, further analysis was carried out using Tukey’s honestly significant difference (HSD) test to explore specific group discrepancies at p < 0.05.
Results
Biochemical parameters in amniotic fluid
Table 1 presents the impact of MP exposure on biochemical parameters in the amniotic fluid of chick embryos, detailing significant changes with a dose-dependent response. Upon comparison using Tukey’s HSD test at p < 0.05, there were statistically significant increases in the levels of urea, creatinine, ALP, AST, and ALT across the groups exposed to MPs relative to the control group. The urea levels in the control group averaged 8.26 ± 1.158 mg/dl, which significantly increased to 16.79 ± 1.569 and 20.35 ± 1.948 mg/dl in groups A (150 µg/ml MPs) and B (300 µg/ml MPs), respectively. Creatinine, ALP, AST, and ALT also exhibited significant elevations in a similar pattern, with group B displaying the highest levels, indicative of a robust dose-response relationship.
Table 1.
Comparison of biochemical parameters in amniotic fluid of chicken embryos: control versus treated groups on day 18 (n=5).
| Biochemical parameters | Control group (C) | Treatment group (A) | Treatment group (B) | p-value |
|---|---|---|---|---|
| Urea (mg/dl) | 8.2604 ± 1.158a | 16.7876 ± 1.569b | 20.3492 ± 1.9481c | 0.00001 |
| Creatinine (mg/dl) | 0.621 ± 0.0871a | 0.7698 ± 0.0812b | 0.848 ± 0.0514c | 0.00144 |
| ALP (IU/l) | 7.2196 ± 1.305a | 270.4562 ± 10.7282b | 292.6852 ± 4.6987c | 0.00001 |
| AST (IU/l) | 9.824 ± 0.7046a | 163.626 ± 6.2069b | 187.5172 ± 34.3834c | 0.00001 |
| ALT (IU/l) | 3.9866 ± 0.4179a | 54.181 ± 1.4794b | 56.822 ± 1.5231c | 0.00001 |
Values are shown as Mean±SD (n=5). Means within the same row for each parameter that carry different superscripts (a,b,c) differ significantly at p < 0.05.
Abbreviations: ALP, alkaline phosphatase; ALT, alanine aminotransferase; AST, aspartate aminotransferase.
Effects on caecal tonsil development
Observations of 18-day-old chick embryos from the control group, depicted in Figure 1 C/a under 10× magnification (H&E stain), reveal a dense accumulation of lymphocytes within the lamina propria, as indicated by the scale bar (160 µm). This observation is corroborated by Figure 1 C/b, which, under 100× magnification (H&E stain, scale bar: 16 µm), confirms the high lymphocyte density in the control group. Conversely, Figure 1 A/a (10× magnification, H&E stain, scale bar: 160 µm) demonstrates a reduction in lymphocyte count by 150 µg/ml in the treatment groups. Similarly, Fig. A/b (100× magnification, H&E stain, scale bar: 16 µm) illustrates a decreased lymphocyte presence in the lamina propria of the treatment group. This reduction is more pronounced in Fig. 1 B/a (10× magnification, H&E stain, scale bar: 160 µm), where the lymphocyte count is significantly lower (300 µg/ml) compared to both A and the control group. This finding is consistent with Fig. 1 B/b (100× magnification, H&E stain, scale bar: 16 µm), which shows the lamina propria of the 300 µg/ml treatment group containing fewer lymphocytes than those in the A and control groups. Table 2 presents a quantitative analysis of lymphocyte counts in the caecal tonsils of chick embryos exposed to different concentrations of MPs. Using Tukey’s HSD test at p < 0.05, the data reveal that the control group (C) had a significantly higher lymphocyte count (237 ± 11.6619) compared to the MP-treated groups. Group A (150 µg/ml MPs) showed a lymphocyte count of 202.8 ± 11.3886, and Group B (300 µg/ml MPs) recorded even lower at 162.8 ± 10.4259. These results indicate a clear dose-response effect, with statistically significant decreases in lymphocyte numbers as MP dosage increased. The differences among the groups underline the pronounced impact of MPs on lymphocyte counts in the caecal tonsils.
Table 2.
Comparison of lymphocyte counts in the caecal tonsils among control (C) and treated (A and B) groups.
| Group | Control (C) | Treatment group (A) | Treatment group (B) | p value |
|---|---|---|---|---|
| Number of lymphocytes | 237 ± 11.6619a | 202.8 ± 11.3886b | 162.8 ± 10.4259c | 0.00001 |
Values are shown as mean ± SD (n=5). Means within the same row for each parameter that carry different superscripts (a,b,c) differ significantly at p < 0.05.
Figure 1.
Histological comparison of caecal tonsils in chick embryos at 18 days of development. The images present sections stained with H&E at different magnifications: 10× (scale bar: 160 µm) and 100× (scale bar: 16 µm). Panels C/a and C/b show samples from the control group, panels A/a and A/b are from the group treated with 150 µg/ml, and panels B/a and B/b are from the group treated with 300 µg/ml. The density of lymphocytes in the lamina propria is emphasized with rectangles, and individual lymphocytes are indicated by circles.
Effects on bursa of Fabricius
In this histological examination of the bursa of Fabricius in 18-day-old chick embryos, the average mucosal fold (MF) count was consistently found at 11 across all observed groups. Delving into follicular details within each fold, no significant discrepancies were detected, as outlined in Table 3. Illustrative micrographs from the control group (Fig. 2C/a and C/b), stained with H&E and viewed at 10× (160 µm) and 100× (16 µm) magnifications, respectively, exhibit a prominent congregation of lymphocytes within lymphatic follicles, evident from circled regions. Upon closer inspection, square markers pinpoint escalated lymphocyte presence within these follicles. Conversely, the group subjected to a 150 µg/ml concentration (Fig. 2A/a, stained with H&E, 10× magnification, 160 µm scale bar) showcased a diminished lymphocyte density within lymphatic follicles compared to the control, highlighted by circular markers. This trend persisted under higher magnification (Fig. 2A/b, stained with H&E, 100×, 16 µm), elucidating a conspicuous reduction in lymphocyte numbers relative to the control.
Table 3.
Comparing MF counts in the Bursa of fabricius between control (C) and treated (A and B) groups.
| Group | Control (C) | Treatment group (A) | Treatment group (B) | p value |
|---|---|---|---|---|
| Number of MF per bursa of Fabricius | 11.4 ± 0.5477a | 11.6 ± 0.5477a | 11.6 ± 0.8944a | 0.86833 |
| Number of lymphatic follicles per MF | 84.6 ± 3.1305a | 79.4 ± 2.4083b | 70.6 ± 2.3022c | 0.00001 |
| Number of lymphocytes per slide | 256.8 ± 5.8052a | 249.2 ± 5.1672b | 223 ± 12.6095c | 0.000108 |
Values are shown as Mean ± SD (n=5). Means within the same row for each parameter that carry different superscripts differ (a,b,c) differ significantly at p < 0.05.
Figure 2.
Histological analysis of the bursa of Fabricius in 18-Day chick embryos: panel A depicts the lymphatic follicles of embryos treated with 150 µg/ml MPs (A/a: H&E stain, magnification 10×, scale bar 160 µm; A/b: H&E stain, magnification 100×, scale bar 16 µm). Panel B presents the follicles in embryos treated with 300 µg/ml MPs (B/a: H&E stain, magnification 10×, scale bar 160 µm; B/b: H&E stain, magnification 100×, scale bar 16 µm). Panel C shows the control group’s follicles (C/a: H&E stain, magnification 10×, scale bar 160 µm; C/b: H&E stain, magnification 100×, scale bar 16 µm). Rectangles indicate the areas with dense lymphocyte populations, while circles mark individual lymphocytes.
Further scrutiny of the group administered a 300 µg/ml concentration (B/a) and stained with H&E at 10× magnification unveiled a notable decrease in lymphocyte concentration within lymphatic follicles, as indicated by circular markers. This observation was corroborated by high-magnification micrographs (B/b, stained with H&E, 100×, 16 µm), demonstrating fewer lymphocytes in lymphatic follicles compared to both the A group and the control.
Table 3 provides insights into the effects of MP exposure on the structural features within the bursa of Fabricius of chick embryos. Statistical analysis using Tukey’s HSD test at p < 0.05 indicated no significant differences in the number of MFs between the control group (11.4 ± 0.5477) and the treatment groups A and B (both recorded 11.6 ± 0.5477 and 11.6 ± 0.8944, respectively), showing stability in this structural parameter across varying MP concentrations. However, significant differences were observed in the counts of lymphatic follicles and lymphocytes among the groups. The control group exhibited the highest count of lymphatic follicles (84.6 ± 3.1305), significantly higher than that observed in treatment group B (70.6 ± 2.3022), suggesting a reduction with increased MP dosage. Similarly, lymphocyte counts were highest in the control group (256.8 ± 5.8052) and decreased in treatment group B (223 ± 12.6095), illustrating a clear dose-dependent effect. These results highlight significant variations in the lymphatic follicle and lymphocyte counts in response to MP exposure, with other structural parameters such as MFs remaining unaffected.
Effects on immunity genes
On the 18th day of the incubation period, we conducted a qPCR test to evaluate the expression levels of the immunity genes AvBD9, AvBD10, and IL6. In comparison to the control group (C), both treated groups (A and B) displayed a decrease in mRNA expression of these immunity genes. Additionally, the expression of AvBD9, AvBD10, and IL6 decreased in a dose-dependent fashion within the treated groups when compared to the control group. This was confirmed through qPCR analysis performed on day 18 of incubation (Fig. 3).
Figure 3.
Expression levels of the immunogenic genes AvBD9, AvBD10, and IL6 in chicken embryos post-titanium dioxide exposure. Notably, groups A and B exhibit significant reductions in gene expression compared to control group C (n=3 per group). ANOVA statistical analysis (p < 0.05) confirms significant downregulation, highlighting the immunomodulatory potential of Titanium dioxide exposure.
Discussions
During this 18-day investigation, both the control and treated groups were closely monitored for any adverse effects. We observed no deaths or harmful outcomes related to the incubation conditions in either group. The selection of MP concentrations for the experiment was not solely influenced by the typical human intake of 0.1 to 5 µg per week, as noted by Senathirajah et al. [19]. It also considered other important factors, such as the possibility of bioaccumulation and the greater biological effects caused by the increased surface area of smaller MP particles. It is essential to consider these exposures in light of various avian species’ body weight and dietary habits to assess the environmental significance and potential health hazards. The present research investigates the effects of MPs on bird species, emphasizing their impact during the embryonic development phase. This focus aids in assessing the environmental and health consequences of MPs on avian populations. Moreover, this study distinctively examines how MPs alter the embryonic microenvironment by analyzing changes in the biochemical composition of amniotic fluid and the expression of immune-related genes.
The findings from this study align with previous research, demonstrating that exposure to MPs significantly influences the biochemical composition of amniotic fluid in chicken embryos. Notable increases in the levels of urea, creatinine, ALP, AST, and ALT were observed in MP-treated groups. These results are consistent with Banaee et al. [20], who found similar elevations in urea, Creatinin, ALT, ALP, and AST levels in pond turtle (Emys orbicularis) exposed to MPs, suggesting a disturbance in metabolic processes and increased oxidative stress, particularly affecting liver and kidney function. The elevated biochemical markers identified in our study could be indicative of hepatotoxicity during embryonic development. The liver and kidneys play essential roles in metabolic regulation and detoxification; thus, the significant changes observed in these biochemical parameters could imply detrimental effects on the physiological development of the embryos. This is further supported by Jin et al. [21], who noted similar biochemical disruptions in mice. Moreover, the dose-dependent increase in these markers highlights the potential for MPs to cause progressive cellular damage and metabolic disturbances. These findings emphasize the need for further investigation into how MPs disrupt biochemical pathways and the broader implications for embryonic health and development.
The results of this study indicate that exposure to MPs leads to significant changes in the immune-related tissues of chicken embryos, specifically within the caecal tonsil and the bursa of Fabricius. These findings are important as they highlight the potential immunotoxic effects of MPs during critical periods of immune system development. In the caecal tonsil, the observed reduction in lymphocyte counts within the lamina propria in treatment groups A and B compared to the control group suggests that MPs may impair lymphocyte proliferation or survival. Lymphocytes are critical for the adaptive immune response, and their decrease could result in compromised immune function, making the embryos more susceptible to infections and diseases. This reduction could be due to direct cytotoxic effects of MPs or due to indirect mechanisms such as oxidative stress or inflammation induced by MPs, as suggested by similar studies on other species. For instance, a study by Huang et al. [22] on the effects of MPs on the immune system of mice also reported reduced lymphocyte proliferation and increased signs of oxidative stress in lymphoid organs. A review study by Yang et al. [23] showed that MPs exposure resulted in alterations in the immunity. These studies, along with the current findings, suggest a broader, potentially cross-species impact of MPs on the lymphatic system and immune health. Furthermore, the alteration in the counts of lymphatic follicles within the bursa of Fabricius, which is vital for B-cell maturation in birds, indicates that MPs might also affect the development of the adaptive immune system more broadly. This could lead to long-term consequences for immune competency and disease resistance in birds.
The findings of the present study demonstrate a reduction in the expression of AvBD9, AvBD10, and IL6 mRNA in experimental groups compared to control groups, indicating potential immunomodulatory effects of MPs exposure on chicken embryos. Yang et al. [23] provide insights into the mechanisms underlying the immunotoxicity of MPs. MPs have been shown to disrupt intracellular signaling pathways, leading to alterations in immune homeostasis and tissue damage. This disruption often involves the generation of ROS, which can trigger inflammatory responses through toll-like receptors (TLRs) and cytokine production. Additionally, the formation of protein-corona complexes on MPs can enhance their toxicity and exacerbate immune responses. These mechanisms align with the observed down-regulation of immune-related genes in the present study, suggesting a potential link between MPs exposure and impaired immune function. A study by Limonta et al. [24] supports the notion that MPs can affect immune system gene expression and epithelial integrity. Transcriptomic analysis revealed alterations in immune-related genes, which could compromise the organism’s defense against pathogens. This aligns with the reduced expression of immune-related genes observed in the present study and underscores the potential consequences of MPs exposure on immune response efficiency. Furthermore, Huang et al. [22], Bacolod et al. 2017 [25], and Sussarellu et al. [26] provide additional insights into the impact of MPs on immune function. Their findings suggest that MPs disrupt immune-related receptors and cell signaling pathways, leading to suppressed immune responses and genotoxicity. This aligns with the observed reduction in immune gene expression in the present study and highlights the potential mechanisms through which MPs exert their immunomodulatory effects, including the accumulation of MPs in the extracellular matrix (ECM), leading to ECM dysfunction and immune receptor suppression. Collectively, these studies provide compelling evidence that exposure to MPs can disrupt immune function through various mechanisms, including oxidative stress, inflammation, and interference with cell signaling pathways. The observed reduction in the expression of immune-related genes in chicken embryos exposed to MPs underscores the need for further research to understand the full extent of immunotoxicity associated with MPs exposure and its implications for embryonic development and health.
Conclusion
The study on the effects of MPs exposure on chick embryos highlights significant immunomodulatory and biochemical alterations. Findings revealed elevated levels of urea, creatinine, ALP, AST, and ALT, indicating disturbances in metabolic processes and potential hepatotoxicity. Additionally, reduced lymphocyte counts in the caecal tonsil and altered immune gene expression underscored immunotoxic effects, compromising immune function and disease resistance in birds. Further research is needed to understand immunotoxicity extent and implications for embryonic health. Addressing these concerns can aid in protecting avian populations and ecosystems from the detrimental impacts of MPs pollution, benefiting global environmental and public health initiatives.
Acknowledgments
The authors would like to express their heartfelt appreciation for the assistance provided by the Institute of Research and Training, Hajee Mohammad Danesh Science and Technology University, situated in Dinajpur-5200, Bangladesh.
Conflict of interest
The authors have confirmed the absence of any potential conflicts of interest.
Author contributions
Md. Sadequl Islam: contributed significantly to conceptualization, investigation, methodology, and validation, as well as writing the original draft and reviewing and editing the manuscript. Md. Ahsanul Kabir: data analysis, review, and writing the manuscript. Sheikh Ismail Ahmed: conducted the experiment, data analysis, review, and writing the manuscript.
